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Increased sensory input from maternal care attenuates neuroendocrine and behavioral responses to stress long-term and results in a life-long phenotype of resilience to depression and improved cognitive function. Whereas the mechanisms of this clinically important effect remain unclear, the early, persistent suppression of the expression of the stress neuro-hormone corticotropin releasing hormone (CRH) in hypothalamic neurons has been implicated as a key aspect of this experience-induced neuroplasticity. Here we tested if the innervation of hypothalamic CRH neurons of rat pups that received augmented maternal care was altered in a manner that might promote the suppression of CRH expression, and studied the cellular mechanisms underlying this suppression. We found that the number of excitatory synapses and the frequency of miniature excitatory synaptic currents onto CRH-neurons were reduced in ‘care-augmented’ rats compared with controls, as were the levels of the glutamate vesicular transporter vGlut2. In contrast, analogous parameters of inhibitory synapses were unchanged. Levels of the transcriptional repressor, neuron-restrictive silencer factor (NRSF), which negatively regulates Crh gene transcription, were markedly elevated in care-augmented rats, and chromatin immunoprecipitation demonstrated that this repressor was bound to a cognate element (NRSE) on the Crh gene. Whereas the reduced excitatory innervation of CRH-expressing neurons dissipated by adulthood, increased NRSF levels and repression of CRH expression persisted, suggesting that augmented early-life experience re-programs Crh gene expression via mechanisms involving transcriptional repression by NRSF.
Early-life experience induces persistent neuroplasticity of the neuroendocrine stress system, with implications for emotional and cognitive function (Levine, 1967). This plasticity, characterized by reduced stress responses (Plotsky and Meaney, 1993; Avishai-Eliner et al., 2001a), increased resilience to depressive-like behavior (Meaney et al., 1991) and improved learning and memory (Liu et al., 2000; Fenoglio et al., 2005), can be induced by brief daily separation of rat pups from the dam during the first weeks of life, which promotes maternal-derived sensory input (Liu et al., 1997; Fenoglio et al., 2006).
At a molecular level, adult rats experiencing augmented maternal sensory input have altered basal expression of key neuronal genes involved in regulating neuroendocrine and behavioral stress responses (Joels and Baram, 2009). These include reduced expression of corticotropin releasing hormone (CRH) in hypothalamic paraventricular nucleus neurons, attenuated hormonal stress response and increased expression of glucocorticoid receptor in hippocampal neurons (Plotsky and Meaney, 1993; Liu et al., 1997; Fenoglio et al., 2005). Experience-induced reduction of CRH expression was apparent already by P9, preceding both the diminished hormonal stress responses and the increased hippocampal GR expression (Avishai-Eliner et al., 2001a). These observations, together with the fact that the ‘experience-augmented’ phenotype could be reproduced by reducing interactions of CRH with its receptor during development (Fenoglio et al., 2005) suggested that reduced CRH expression was an early, key element of the process leading from enhanced maternal sensory input to the enduring phenotype described above.
What is the nature of signals, derived from maternal sensory experience that reaches CRH neurons? The CRH neuron is a component of a neuronal network activated by maternal care (Fenoglio et al., 2006). Therefore, it is reasonable to assume that augmented early-life experience alters excitatory and / or inhibitory synaptic input onto CRH neurons, and that this alteration, in turn, triggers molecular machinery that enduringly reduces CRH expression.
Synaptic innervation of individual (including hypothalamic) neurons has recently emerged as dynamic and modulated by experience (Verkuyl et al., 2004; Horvath, 2005). For CRH neurons, the majority of input is mediated by GABAergic and glutamatergic synapses (Boudaba et al., 1997; Miklos and Kovacs, 2002; Ziegler et al., 2005), via GABAA (Cullinan, 2000) and glutamate receptors (Aubry et al., 1996; Kiss et al., 1996; Di et al., 2003) GABA and Glutamate are transported into presynaptic vesicles by vesicular transporters: vGat (McIntire et al., 1997) and vGlut (Herzog et al., 2001), respectively, so that these transporters provide specific markers for inhibitory and excitatory afferent synaptic contacts onto CRH neurons. Here we combined quantification of these markers, using western blot, quantitative confocal and electron microscopy, and electrophysiology, to examine the effects of early-life experience on excitatory and inhibitory synapses abutting CRH neurons. We then examined the molecular machinery underlying the persistent suppression of CRH expression, testing if augmented early-life experience increased expression of the repressor neuron-restrictive silencing factor (NRSF; Mori et al., 1992; Palm et al., 1998), because of the presence of a functional binding site for this repressor within the Crh gene (Seth and Majzoub, 2001).
Thirty-two, timed-pregnant Sprague-Dawley rats were housed in an uncrowded, quiet animal facility room on a 12 h light/dark cycle and were provided with food and water ad libitum. Parturition was checked daily, and the day of birth was considered postnatal day (P) 0. On P2, all pups were briefly gathered, and 10 pups assigned at random to each dam. They experienced one of the following early-life rearing conditions: (a) Brief separation from the dam (augmented care, or augmented-experience'), which took place daily from P2 to P8. (b) Undisturbed controls, that remained in cages that were not touched between P2 to P8. Rats were either sacrificed on P9, between 08:00 and 12:00 or maintained under standard animal facility rearing conditions, weaned on P21, and grown, 3-4-per cage into adulthood. All experiments were approved by the University Animal Care Committee and conformed to National Institutes of Health guidelines.
Cages were brought into laboratory daily at 8:30 AM. The dam and the pups were placed into separate bedded cages (pups were kept euthermic via a heating pad located underneath the cage). After 15 min, pups were placed back into their home cage, followed by the dam, and returned to the vivarium. Undisturbed litters remained in the vivarium from P2 to P9. For all experimental groups, cage change did not occur during this time. After pups and were returned to the vivarium, maternal behavior, (including licking and grooming, was observed using a protocol based on (Liu et al., 1997) as used by (Fenoglio et al., 2006). Maternal behavior was also determined in undisturbed litters. Each maternal observation session consisted of 10 epochs of 3 min each. Within each epoch, the duration of licking and grooming of pups was recorded during the first min. The total amount of time spent licking and grooming was scored per session, added for all the sessions and multiplied by three to provide the total amount of maternal care during the 30 min session.
Rats were rapidly decapitated at P9 (n=5 for control and n=6 for experience-augmented pups) and between P45 and P60 (n=4 /group), and the brains were quickly dissected and frozen in dry ice. Brains were sectioned at 20 μm using a cryostat, collected onto gelatin coated slides and stored at -80 °C. ISH histochemistry was conducted as described previously using deoxy-oligonucleotide probes (Avishai-Eliner et al., 2001b). Probe was labeled with 35S using routine terminal deoxynucleotide transferase methodology. Briefly, sections were brought to room temperature, air-dried and fixed in fresh 4% buffered paraformaldehyde for 20 min, followed by dehydration and rehydration through graded ethanols. Sections were exposed to 0.25% acetic anhydride and 0.1 M triethanolamine (pH 8) for 8 min and were dehydrated through graded ethanols. Pre-hybridization and hybridization steps were performed in a humidified chamber at 42 °C in a solution of 50% formamide, 5× SET, 0.2% SDS, 5× Denhart's, 0.5 mg/ml salmon sperm DNA, 0.25 mg/ml yeast tRNA, 100 mM dithiothreitol and 10% dextran sulfate. Following a 1 h prehybridization, sections were hybridized overnight with 0.25×106 CPM of labeled probe. After hybridization, sections underwent serial washes at 42 °C, most stringently at 0.3× SSC for 30 min at room temperature. Sections were then dehydrated through increasing ethanol concentrations, air-dried and apposed to film (Kodak BioMax MR Film, Eastman Kodak Co. NY, USA), for 5–10 days.
ISH signal was analyzed on scanned, digitized PVN images of sections at coronal levels corresponding at 3.8-3.5 mm anterior to bregma. The Image Tool software program (UTHSC, San Antonio, TX) was used (Fenoglio et al., 2006) after determining the linear range of optical densities (ODs) using 14C standards. Values were only used if they fell in the linear range and were corrected for background by subtracting the OD of signal over the thalamus. OD from two optimal sections was averaged to generate an expression value for each PVN, which is expressed in nCi/g. These values were then used to calculate group means± standard error of the mean (SEM).
At P9 or at P45 rats were anesthetized with sodium pentobarbital (100 mg/kg, i.p.) and then perfused through ascending aorta with 0.9% saline solution, followed by freshly prepared, cold 4% PFA in 0.1 M sodium phosphate buffer (PB) pH 7.4, cryoprotected in 15% and 30% sucrose/PB solution, and stored at -80°C. Brains were sectioned at 30μm using a cryostat and sections were collected as four series with 90μm intervals in between sections in tissue-culture wells containing 0.1 M PB. Every fourth matched section was subjected to ICC. For each experiment, sections of experience-augmented and control rat brains were processed concurrently in parallel wells. For single labeling CRH ICC, free-floating sections of experience-augmented (n=5) and control (n=5) rats were subjected to standard avidin-biotin complex (ABC) methods (Fenoglio et al., 2006). Briefly, after several washes with 0.01 M phosphate-buffered saline (PBS) containing 0.3% Triton X-100 (pH 7.4, PBS-T), sections were treated for 30 min in 0.3% H2O2 in PBS, followed by blockade of non-specific sites with 1% bovine serum albumin (BSA) in PBS for 30 min and incubation for 48 hours at 4°C with anti-CRH (1:40,000; gift from Dr. W.W. Vale, Salk Institute, La Jolla, CA, USA) in PBS After 3 times 5 min washes in PBS-T, sections were incubated in biotinylated rabbit-anti-goat IgG (1:200, Vector, Burlingame, CA) in PBS containing 1% bovine serum albumin for 2 hours at room temperature. After washing in PBS-T (3 × 5 min), sections were incubated in ABC solution (1:100; Vector) for 2 hours at room temperature. Sections were then rinsed again in PBS (3 × 5 min). The reaction product was visualized by incubating sections for 8-10 min in 0.04% 3,3-diaminobenzidine (DAB) containing 0.01% H2O2.
All counts were performed without knowledge of experimental group (blindly). CRH immunopositive cells were visualized using a Nikon E400 microscope, and counted in anatomically matched sections using systematic sampling methods: Briefly, CRH immunopositive cells were counted in four sections which were 90μm apart per animal. We obviated double counting by focusing throughout the 30μm section and including a cell only at the level where it had a clear complete nuclear profile. Because the diameter of the CRH positive cell nucleus was considerably smaller than the thickness of each cryostat section, each cell was counted once only. The PVN and ACE were sampled at coronal levels 3.8-3.5 mm anterior to bregma, and the BnST at levels corresponding to 5.0-4.7mm anterior to bregma. The intensity of CRH immunoreactivity was analyzed on photomicrographs taken through a digital camera (Spot Digital camera, RT color V3.0, Diagnostics Instruments MI, USA) with standardized light source and standardized exposure. Analyses were carried out using ImageJ (version 1.41, NIH, Bethesda, MD, USA). Densities are expressed in OD units after correcting for background by subtracting the density of the immunoreactive signal over the anterior commissure. Numbers of CRH immunopositive neurons and CRH OD from four sections were averaged. The numbers of cells/section and OD/section were used to calculate group means ± SEM.
For double labeling ICC of CRH and vGlut2 (the predominant vGlut isoform in the hypothalamus; Herzog et al., 2001), sections of experience-augmented and control rats (n = 3/group) were washed 3×10 min in 0.01 M PBS, and treated with 1% Triton X-100 in 0.01 M PBS, for 30 min. Sections were placed in 2% normal goat serum and incubated for 48 hours at 4°C in a mixture of rabbit anti-CRH serum (1:10,000) and guinea pig anti-VGlut2 serum (1:10,000; AB5907; Chemicon). After three, 10 min washes in 0.01 M PBS, sections were incubated in a secondary antiserum cocktail (1:400; goat-anti-rabbit IgG 568 and goat-anti-guinea pig IgG Alexa Fluor 488; Molecular probes, Eugene, Orgeon).
The number of vGlut2 boutons contacting PVN-CRH neurons was assessed by confocal laser-scanning microscopic analysis (n=66 cells for control and 76 cells for experience-augmented rats). Images of the parvocellular division of the PVN (at coronal levels 3.8-3.5 mm anterior to bregma) were collected using a Zeiss LSM510 confocal scanning system. Five, 2 μm thick optical sections were collected along the Z-axis throughout the thickness of the whole neuron Images were acquired with excitation wavelength of 488nm (green) and 543nm (red) with a 40× oil objective employing the minimum pinhole size. Images were imported as TIFF files at the resolution of 1024×1024 pixels. vGlut2 boutons contacting PVN-CRH neuron were counted through the complete Z-axis in each optical section, and averaged (vGlut2 boutons/optical section/CRH cell). A vGlut2 bouton was considered to be apposed to the CRH cell body only when there was no visible space between the CRH cell membrane and the bouton. In addition cell size was measured in the central optical section for each CRH neuron using Zeiss LSM Image browser software (version 18.104.22.168). Analyzed cells were selected based on the following criteria: 1) CRH immunopositive; 2) fully visible soma within the Z-stack; 3) a clearly identifiable nucleus. After all of the quantitative analyses were completed, images used for illustration were optimized for brightness and contrast using Adobe Photoshop 7.0.
Samples consisted of micro-punched PVN (Palkovits, 1973) or dissected thalamus from an individual rat. Rats were rapidly decapitated at P9 (n=12 per group) and at P45 (n=5 per group), and the brains were quickly dissected and frozen on powdered dry ice. The PVN was punched out (needle gauge 21) from consecutive 150μm thick cryostat sections. Tissue was homogenized in 1.5 ml eppendorf tubes in ice cold 0.32 M sucrose, 0.1 M Tris–HCl (pH 7.4) containing a protease inhibitor cocktail (PIC, Complete™; diluted according to manufacturer's instructions; Roche, Alameda, CA). Protein concentration was determined (Bio-Rad, Hercules, CA), and equal amounts of protein were diluted in Laemmli buffer, separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE), and visualized using the enhanced chemiluminescence (ECL)-Plus kit (Amersham Pharmacia Biotech, Piscataway, NJ) as previously described (Brewster et al., 2005). Briefly, 2 μg of protein extracts were separated on a 10% SDS-PAGE and transferred to Hybond-P polyvinyl difluoride membranes (Amersham Pharmacia Biotech). Membranes were blocked with 10% nonfat milk in 1× PBS overnight at 4 °C and probed with guinea pig anti-vGlut2 (1:800,000, AB5907; Chemicon, Temecula, CA), or rabbit anti-vGat (1:150,000 generous gift of Dr. Edwards UCSF, San Francisco, CA, USA), or rabbit anti-NRSF (1:20,000, H-290-sc25398; Santa Cruz Biotechnology, Santa Cruz, CA, USA) in each case combined with rabbit anti-actin (1:60 000; Sigma, St Louis, MO). Following washes in PBS-1% Tween (PBS-T) (3 × 5 min), membranes were incubated with secondary antibodies made in donkey, conjugated to horseradish peroxidase (1:10 000) in 5% non-fat milk in PBS for 1 h at room temperature. Membranes were washed in PBS-T (3 × 5 min) and incubated with ECL-Plus. Immunoreactive bands were visualized by apposing membranes to Hyperfilm™ ECL (Amersham Pharmacia Biotech). PVN and thalamic extracts of individual rats of different groups were run concurrently on the same gel. Specificity of signal was examined by excluding the primary antibodies in the presence of the secondary antibodies. These treatments resulted in no immunoreactive-vGlut2, vGat or NRSF bands.
Western blot data acquisition and analysis were accomplished by measuring OD of immunoreactive bands from the ECL films using ImageTool. Several durations of exposure were carried out, to ascertain that optical densities were in a linear range. The OD of vGlut2, vGat and NRSF bands was normalized to β-actin of the same sample. Individual OD's were normalized to the mean value of the control OD and used to generate mean values / group ± SEM.
P9 Rats were anesthetized with sodium pentobarbital (100 mg/kg, i.p.) and then perfused through the ascending aorta with 0.9% saline solution, followed by freshly prepared, cold 4% PFA; 15% picric acid and 0.08% glutaraldehyde in 0.1 M PB, pH 7.4. Brains were removed and postfixed in 4% PFA overnight and stored in 0.1M PB. Brains were sectioned at 30μm using a vibratome (Lancer) and sections were collected in multi-well plates containing 0.1 M PB. For CRH ICC, free-floating sections of experience-augmented (n=5) and control (n=5) rats were processed according to standard avidin-biotin complex (ABC) methods with minor adaptations for the electron microscopic staining. Specifically, sections were treated for 30 min in 0.3% H2O2 in PBS. Sections were incubated for 1 hour in freezing solution (0.1M PB, 25% sucrose; 10% glycerol) followed by freeze/thaw in liquid nitrogen repeated 3 times. Then, sections were washed several times with 0.01 M PBS followed by incubation in 1% BSA in PBS for 30 min to block non-specific staining. After a 10-minute rinse in PBS, sections were incubated for 48 hours at 4°C with anti-CRH (1:40,000) in PBS containing 1% bovine serum albumin and washed 3 times in PBS-T, 5 min each. Subsequently, sections were incubated in biotinylated rabbit-anti-goat IgG (1:200, Vector, Burlingame, CA) in PBS for 2 hours at room temperature. After washing in PBS (3 × 5 min), sections were incubated in ABC solution (1:100; Vector) for 2 hours at room temperature. Sections were then rinsed again in PBS (3 × 5 min). The reaction product was visualized by incubating sections for 8-10 min in 0.04% 3,3-diaminobenzidine (DAB) containing 0.01% H2O2. Sections were then osmicated (1% OsO4 in PB) for 30 min, dehydrated through increasing ethanol concentrations (using 1% uranyl acetate in the 70% ethanol for 30 min), and flat embedded in Durcupan between liquid release-coated slides and coverslips (Electron Microscopy Sciences, Fort Washington, PA) followed by capsule embedding. Blocks were trimmed and ribbons of serial ultrathin sections were cut (using a Leica Ultracut E), collected on Formvar-coated single slot grids and examined under an FEI Biotwin electron microscope.
The quantitative and qualitative analysis of synapse number was performed in a blinded fashion. To obtain a complementary measure of axosomatic synaptic number, unbiased for possible changes in synaptic size, the dissector technique was used. On consecutive 90-nm-thick sections we determined the average projected height of the synapses and used about 30% of this value as the distance between the dissectors. On the basis of this calculation, the number of axosomatic synapses was counted in 2 consecutive serial sections about 270 nm apart (termed reference and look-up sections) of 10 CRH-immunolabeled perikarya profiles in each animal. Synapse characterization was performed at a magnification of 20,000. Symmetric and asymmetric synapses were counted on all selected neurons only if the pre- and/or postsynaptic membrane specializations were seen and synaptic vesicles were present in the presynaptic bouton. Synapses with neither clearly symmetric nor asymmetric membrane specializations were excluded from the assessment. The plasma membranes of selected cells were outlined on photomicrographs, and their length was measured with the help of a cartographic wheel. Plasma membrane length values measured in the individual animals were added, and the total length was corrected to the magnification applied. Synaptic densities were evaluated according to the formula NV = Q−/Vdis, where NV represents number per volume and Q– represents the number of synapses present in the reference section that disappeared in the look-up section and Vdis is the dissector volume (volume of reference), the area of the perikarya profile multiplied by the distance between the upper faces of the reference and look-up sections (i.e., the data are expressed as numbers of synaptic contacts per unit volume of perikaryon). Section thickness was determined using the Small's minimal fold method. The synaptic counts were expressed as numbers of synapses on a 100μm membrane length unit.
Whole cell recordings were made from presumed CRH neurons from the parvocellular paraventricular hypothalamic nucleus. Rats were rapidly decapitated at P9 (n=8 rats per group), and at P30 (n=3 rats per group) brains quickly dissected and 300μm hypothalamic slices cut using a vibratome. Hypothalamic slices were maintained at 33 °C and perfused with ACSF. The bath solution (ACSF) consisted of 124mM NaCl; 3mM KCl; 2mM CaCl2; 2mM MgCl2; 1.23mM NaH2PO4; 26mM NaHCO3; 10mM glucose pH=7.4 with NaOH and was continuously bubbled with 5% CO2 and 95% O2. The patch pipettes were made of borosilicate glass (World Precision Instruments) with a Sutter micropipette puller (PP-97). The tip resistance of the recording pipettes was 4-6 MΩ after filling with a pipette solution containing 145mM KMeSO4, 1mM MgCl2; 10mM Hepes; 1.1 mM EGTA; 2mM Mg-ATP; 0.5 mMNa2-GTP pH=7.3 with KOH and 0.1% Lucifer Yellow (LY). After a gigaohm seal and a whole cell access were achieved, the series resistance was between 20 and 40 MΩ and partially compensated by the amplifier. Both input resistance and series resistance were monitored throughout the experiments. mEPSCs and mIPCSs were recorded under voltage clamp in the presence of TTX and bicuculline, with pipette solution containing KMeSO4 (for mEPSCs) or TTX and CNQX plus AP-5, with pipette solution containing KCl (for mIPSCs) with a multiclamp 700A amplifier (Axon Instruments, Inc). The use of KCl significantly increases the conductance of GABA-A receptors and makes recording a mIPSCs easier. PVN cells were held at −60 mV. Detection of mEPSC and mIPSC events were performed offline with the software Axograph 4.9 (Axon Instruments, Inc.). Frequency and Amplitude of mEPSCs (or mIPSCs) were generated after detection of mEPSC events, as described previously (Gao and van den Pol, 2001). Putative CRH neurosecretory parvocellular PVN neurons were identified on the basis of their visualized position within the PVN and based on the shape of the neuron.
After recording with electrodes containing LY in the pipette solution, slices were postfixed overnight in 4% paraformaldehyde at 4 °C overnight, then washed in 0.1 M PB, embedded in agar and sliced with vibratome to 50 um thick sections. After washes in PBS-T, sections were incubated with rabbit anti-CRH (1:10,000) in 1% BSA in PBS-T for 48h at 4°C. After 3 5 min washes in PBS-T, goat anti-rabbit antibody (Alexa Fluor 568 1:400) in 1% BSA in PBS-T was applied for 3 h to label CRH neurons. Sections were then sealed on glass slides with Vectashield to avoid bleaching, then examined with a fluorescent microscope (Zeiss Axiophot microscope). Overlap of LY images with Alexa Fluor 568 staining was taken as identification of CRH neurons; however leakage of cell contents including the LY in the slices impeded positive identification of the recorded neurons. Therefore, we considered for analysis only recorded neurons residing in the CRH-rich, dorsomedial parvocellular field of the PVN.
Chromatin immunoprecipitation (ChIP) assay was based on a fast ChIP protocol (Nelson et al., 2006). Brains were dissected from P12 rats, hippocampi isolated and frozen in tubes on dry ice, and stored at -80°C until use. All ChIP buffer solutions contained Protease inhibitor complex (Roche). Cross linking was achieved by adding 1 ml 1% formaldehyde in PBS, dicing the tissue with scissors, and incubating at room temperature for 20 min. Samples were spun 5 min at 800g at 4°C and the supernatant discarded. Pellets were washed twice with 1ml 0.125M glycine in PBS, spun 5 min at 800g, and the supernatant discarded. Next, tissue was homogenized in 1ml hypotonic buffer (10mM KCl, 10mM Tris pH 8, 1.5mM MgCl2) using a Dounce homogenizer with an A pestle. Samples were then incubated on ice for 15 min allowing the cells to swell, followed by the addition of 100μL of 10% NP-40 and spinning of samples 1min at 16,000g. The supernatant was discarded and 360μL of RIPA lysis buffer (50mM Tris, 150mM NaCl, 1% NP-40, 0.1% SDS, 5mM EDTA, 1mM EGTA) was added to the remaining nuclear pellet. Samples were then sonicated for 10 min on a high setting (30sec on/30sec off) using a Bioruptor sonicator (Diagenode, Sparta, NJ, USA). Samples were then spun 5 min at 16,000g and the top 350μL of the supernatant was added to 100μL of 50% Protein G Agarose bead slurry blocked with salmon sperm DNA (Upstate, Billerica, MA, USA) and 1mL of RIPA buffer. Samples were incubated on a rotator at 4°C for 1 hour, spun at 3,300g for 1min and transferred to new tubes as ChIP input samples. From ChIP input tubes 425μL samples were aliquoted to new tubes for incubation with 2.5μg of NRSF antibodies (H290, Santa Cruz) or IgG. Samples were incubated on a rotator at 4°C overnight, then spun at 16,000g for 10 min and the top 360μL of the supernatant was transferred to a tube containing 30μL of 50% Protein G Agarose beads. Samples were incubated on the rotator at 4°C for 1 hour and then spun at 3,300g for 1 minute. Supernatant was discarded and the beads were washed 5 times by adding 1mL of RIPA buffer, incubating on the rotator at 4°C for 5 min, spinning at 3,300g, and aspirating the supernatant. To 20μL aliquots of the original ChIP input samples and to each pellet, 65μL of 20% Chelex (Biorad, Hercules, CA, USA) in water was added. The tubes were briefly mixed with a vortex mixer and incubated in boiling water for 10 min. The samples were allowed to cool and spun at 16,000g for 2 min. The top 35μL of supernatant containing immunoprecipitated or input DNA was transferred to new tubes for analysis.
Polymerase Chain Reaction (PCR) on recovered DNA was performed using primers directed at the Crh gene (Forward –AGT TTG GGG AAG AC T TAG GAA GAG, Reverse – CTA TCC GAC AGA CAC AGA CAA GAC) or the Beta Actin gene (Forward – GAC TAC CTC ATG AAG ATC CTG ACC, Reverse – GAG ACT ACA ACT TAC CCA GGA AGG). Gotaq green (Promega, Madison, WI, USA) polymerase mix was used with 10μM of each primer and 1-4μL of DNA template per 20μL reaction. ChIP input DNA samples were used to adjust the amount of DNA used in each PCR to equivalent levels. After 30 cycles of PCR, products were run through gel electrophoresis along with samples of a quantitative DNA ladder (Bioline Hyperladder I). Ethidium bromide and UV light was used to visualize the DNA bands and images were captured. Image analysis was performed on blinded samples using ImageJ (NIH) software to produce a standard curve from the quantitative DNA ladder. Band intensities for the PCR products were quantified using the standard curve.
All analyses were conducted without the knowledge of treatment. The significance of differences between groups was set at 0.05. Differences in the maternal behavior of control and experimental dams were analyzed using repeated measures analysis of variance (RMANOVA), in which treatment and time were included as fixed effects and dam as a repeated (subject) effect, using SAS v.9.2 (SAS Institute Inc., Cary, NC, USA. 2007). In order to correct for potential litter effect, the significance of differences among groups were examined using a mixed model analysis of variance (ANOVA) with treatment as a fixed effect, and the dam as a random effect, using SAS
Brief (15 min) separation of pups from their mother (‘handling’), has been shown to enhance maternal nurturing behaviors, such as licking and grooming, toward the pups (Brown et al., 1977; Liu et al., 1997; Fenoglio et al., 2006). In addition, our previous work found that bursts or vigorous bout of sensory input to the pups was generated during the 30 minutes following the return of the pups to the dam. These findings were confirmed in the current experiments: the return of the pups to the home cage elicited a barrage of maternally derived sensory stimulation. Specifically, maternal licking and grooming was increased two-fold during the first 30 min after the pups' return (304.2±14.4 seconds in vs 614.4±28.2 seconds in controls vs separated dams; F1,31 = 39.94, P < 0.0001) These bursts of sensory stimulation of the pups occurred daily throughout the week of the experiment (Fig. 1).
Hypothalamic parvocellular CRH expression at the mRNA and protein levels were examined on P9 in rats experiencing the augmented maternal care from P2 to P8, compared to controls that were raised without disturbance. CRH mRNA levels in the PVN of separated rats (‘experience-augmented’) were significantly lower than those of control pups (61±6.3 %; P < 0.05; Fig. 2A), and this was observed already by P9, consistent with earlier findings (Avishai-Eliner et al., 2001a; Fenoglio et al., 2006). These changes in mRNA expression were translated into changes in protein expression: as shown in Fig. 2B,C, the intensity of CRH immunoreactivity over the PVN was reduced in the experience augmented group (0.13±0.04 optical density [OD] units/section in experience-augmented vs 0.16±0.05 OD units/section in control animals P<0.05), whereas the total number of CRH-expressing neurons was unchanged (71±5 cells per section for control and 68±5 for experience-augmented rats). The effects of augmented early-life experience were specific to the CRH-expressing neuronal population in the hypothalamic PVN: neither the overall OD nor the total number of CRH immunopositive cells were altered in the central nucleus of the amygdala (ACe) or the bed nucleus of the stria terminalis (BnST). Thus, CRH immunoreactivity in ACe in P9 experience-augmented and control rats averaged 0.06±0.02 and 0.06±0.02 OD units/section, respectively, and the numbers of neurons expressing CRH in ACe averaged 12±1 cells/section and 13±1cells/section respectively. The BnST of control rats had an average of 0.05±0.02 OD units/section and 48±2 CRH expressing neurons/section, compared with 0.05±0.02 OD units/section and 40.0±5 cells/section in BnST from experience-augmented rats.
The longevity of the altered CRH expression was examined in rats experiencing augmented maternal care from P2 to P8 and killed as young adults (at P45-P60). At the mRNA level, PVN CRH expression remained lower in experience-augmented rats (50±4 % of control values P < 0.001; Fig. 2D), consistent with our previous findings (Avishai-Eliner et al., 2001a; Fenoglio et al., 2005). This reduction was evident also at the protein level (Fig. 2E,F): the number of CRH-expressing neurons was lower (58±4 neurons/section) in adult experience-augmented rats compared with the control cohort (75±4 neurons/section; P < 0.05), while the intensity of CRH immunoreactivity was not altered (0.06±0.02 OD units/section in control vs 0.07±0.02 OD units/section in experience-augmented rats). Similar to P9, this reduction was specific to the PVN, because no differences in the number of CRH expressing neurons or intensity of immunoreactive signal were detected in the ACe (0.10±0.05 OD units/section and 47±10 cells/section for control and 0.12±0.05 OD units/section and 44±4 cells/section for experience-augmented rats), or in the BnST (0.13±0.05OD units/section and 17±6 cells/section for control and 0.13±0.07 OD units/section and 12±2 cells/section for experience-augmented rats). Interestingly the numbers of CRH cells in the ACe and BnST differed in both groups between P9 and P45, with inverse trends in cell numbers with age in the two structures. We can only speculate that this might be a result of developmental changes.
The reduction of CRH expression in PVN of experience-augmented rats starts early (P9), endures for life, and is associated with life-long reduction in hormone release in response to stress, potentially indicating altered sensitivity of the CRH neuron to excitation by stress. Therefore, we tested the possibility that augmented early-life experience leads to a reduced excitatory drive (or increased inhibition) onto CRH-expressing neurons.
We first examined whether the levels of the vesicular transporters of glutamate (vGlut2) and GABA (vGat), which are markers of presynaptic elements of glutamatergic and GABAergic synapses, respectively, were influenced by augmented maternal care. In punched PVN tissue, vGlut2 was detected as single major band (~60 kDa; Fremeau et al., 2004). Protein levels of vGlut2 (normalized per actin and per controls), were significantly lower in experience-augmented compared to control rats: 60.5±11.4 vs 101.7±12.5 OD units, respectively (P < 0.01; Fig. 3A,C). The changes in vGlut2 were selective to the PVN, and were not observed in the thalamus (100.0±32.4 in controls vs 92.9±36.9 in experience-augmented; Fig. 3B,C). In contrast to vGlut2, protein levels of vGat were unaffected by early-life experience (49.1±6.6 vs 59.1±8.7 OD units in experience-augmented and control groups; Fig. 3D).
To examine whether the reduction in vGlut2 protein levels in ‘experience-augmented’ rat PVN represented a reduced number of glutamatergic, excitatory synapses onto CRH neurons, we identified excitatory glutamatergic boutons on CRH neurons using double immunolabeling for vGlut2 and CRH. In virtual confocal sections (2μm), vGlut2–ir axon varicosities were frequently juxtaposed onto somata and proximal dendrites of CRH-ir neurons. The CRH-expressing neurons were located in the medial parvocellular subdivision of the PVN (Sawchenko and Swanson, 1985), and their total numbers per section did not differ among the two experimental groups (see above). The numbers of vGlut2 boutons on individual CRH-ir cells were lower in experience-augmented rats (5.4±0.9 vGlut2 boutons /optical section/cell) compared with controls (8.4±0.9; P < 0.05; Fig. 4A-D). The size of CRH neurons, as assessed by measuring cell surface was not affected by the early-life experience (perimeter: 32.87±0.48 μm; n=37 cells from 3 rats, in controls and 33.36±0.51 μm; n=41 cells from 3 rats, in experience-augmented pups; P>0.3).
The reduced number of vGlut2-positive, glutamatergic synapses abutting CRH-expressing cells could be a result of either diminished number of synapses or reduced synapse size. To distinguish between these two possibilities, we examined the density and structural identity of synapses onto CRH cells using electron microscopy. After immunolabeling for CRH, symmetric (inhibitory) and asymmetric (excitatory) synapses onto CRH neurons in the parvocellular PVN were examined using stereological principles. Total synaptic density was lower in experience-augmented rats compared to control cohorts (18.9±6.2 vs 34.8±5.2 synapses/100 μm somatic membrane; P < 0.05; Fig. 5A,B). Further analysis revealed that this reduction was attributable primarily to drastically decreased density of asymmetric, excitatory, synapses (from 11.6±3.7 to 3.4±2.2 synapses/100 μm somatic membrane; P < 0.05; Fig. 5A,B). In contrast to this ~70% reduction of asymmetric synapses, the number of symmetric, inhibitory synapses were not influenced by early-life experience (15.5±5.2 in experience-augmented rats vs 20.9±2.3 synapses/100 μm somatic membrane in undisturbed controls; Fig. 5A,B).
The data above indicated that CRH-expressing neurons in the PVN of experience-augmented rats were contacted by fewer asymmetric, vGlut2-positive excitatory synapses. However, whether these structural changes influenced functional excitatory input impinging on these CRH-neurons was unclear. Therefore, afferent inputs to presumed CRH neurons were measured using whole patch-clamp recordings in acute hypothalamic slices from experience-augmented and control P9 rats. Cells from the CRH-rich dorsal parvocellular region of the PVN (Fig. 6E) were held at −60mV in the whole-cell voltage clamp configuration, in the presence of tetrodotoxin (TTX) to block all action potential-driven PSCs. This enabled us to examine the frequency of miniature excitatory and inhibitory postsynaptic currents (mEPSC/mIPSC) which arise from spontaneous vesicle fusion and typically reflect the number of transmitter release sites (or the probability of release) (Regehr et al., 2001). Analysis revealed that augmented early-life experience drastically reduced the frequency of mEPSC (17.8±2.8/min (n=17) compared to that in controls (44.2±9.4/min (n=15); P < 0.01; Fig. 6A,B), consistent with reduced numbers of presynaptic terminals (or reduced probability of release). Whereas this reduced frequency of mPSC events is indicative of presynaptic changes, the amplitude of these currents is an indication of postsynaptic changes (e.g. receptor number, function or both; Regehr et al., 2001). mEPSC amplitude was modestly increased in the experience-augmented rats (-12.4±0.7 pA, n=15) when compared to the control pups (-10.5±0.6 pA (n=14); P < 0.05; Fig. 6B). In contrast to the changes in excitatory synapses, the frequency of mIPSCs did not differ between groups (47.5±13.6/min; n=15 in control vs 52.8±7.4/min n=16 in experience-augmented rats; Fig. 6C,D), while the amplitude increased modestly (-30.7±2.3 pA (n=15) vs -47.8±6.8 pA (n=16); Fig. 6D). Thus, the electrophysiological data supported the results obtained with quantitative biochemical methods and confocal and electron microscopy.
CRH expression is repressed for life by augmented early-life experience (Fig. 2). The data above raised the possibility that this repression was both initiated and maintained by reduction of excitatory input. Alternatively, repression of CRH expression after augmented early-life experience might become autonomous, so that altered innervation of the CRH neuron was not required for the maintenance of this effect. To address this question, we examined excitatory and inhibitory synaptic input onto hypothalamic CRH-expressing neurons in older rats (P30-45). Examining the levels of vGlut2, we found that by P45 protein levels of vGlut2 were no longer significantly lower than those of the control rats (145.9±19.91 vs 112.8±47.86 OD units (normalized per actin) P > 0.05), consistent with normalization of excitatory synapse input onto CRH neurons of the former. In accordance with this notion, the frequency of mEPSC and mIPSC of presumed CRH neurons in the PVN in experience-augmented and undisturbed control P30 rats were no longer different: Frequency of mEPSC: 193±31/min; n=15 in control vs 293±48/min n=20 in experience-augmented rats; frequency of mIPSC: 276±105/min; n=6 in control vs 414/±83 min n=6 in experience-augmented rats; P > 0.05 for all comparisons. Taken together, these data indicate that whereas diminished excitatory input, resulting from reduced numbers of excitatory synapses onto CRH neurons of experience-augmented rats, might be involved in initiating the molecular machinery that represses CRH gene expression in these cells, reduced excitation was not necessary to maintain the life-long repression of the Crh gene.
If experience-induced ‘re-wiring’ of CRH-expressing neurons is not required for maintaining the repression of the Crh gene, what might the responsible molecular mechanisms for this persistent repression be? To address this question we evaluated the role of NRSF in regulating the Crh gene by testing the ability of NRSF to bind to the Crh gene and by measuring the effects of augmented early-life experience on levels of NRSF in hypothalamic neurons. Using chromatin immunoprecipitation followed by quantitative PCR, we found that amount of ‘Crh-gene’ DNA amplified by PCR from chromatin immunoprecipitated with antiserum to NRSF was 39.2±4.7ng, indicating binding of the repressor to the regulatory region of the gene (Fig. 7A,B). The amount of Crh-gene DNA immunoprecipitated with non-immune IgG and amplified by PCR were undetectable under the ChIP conditions we used, as was the amount of amplified PCR product from the anti-NRSF immunoprecipitated DNA of the intronic region of actin, a gene devoid of NRSE sites (Fig. 7A,B) Quantifying NRSF levels in punched PVN tissue by western blot analysis showed that NRSF migrated as a single major band (~160 kDa). Protein levels of NRSF (normalized per actin), on P9 were dramatically higher in experience-augmented compared to control rats: 147.6±23.6 vs 28.0±7.5 OD NRSF/actin, respectively (P < 0.05; Fig. 7C,E). The experience-induced upregulation of NRSF persisted long term: NRSF levels were still strikingly higher in young-adult experience-augmented rats compared to controls (655.5±332.9 vs 41.6±10.8 OD NRSF/actin; P < 0.05; Fig. 7D,E). This augmentation of NRSF expression was selective to the PVN, because no difference in NRSF levels was found in the thalamus (199.8±47.1 and 195.2± 46.9 OD NRSF/actin in controls and experience-augmented rats respectively; Fig. 7F). These results indicate that upregulation of NRSF levels likely contributes to both initiation and persistence of the repressed CRH expression after augmented early-life experience.
The studies described here demonstrate that following augmented early-life experience: (1) Expression of CRH mRNA and protein in hypothalamic PVN neurons is depressed, and (2) this depression is maintained to adulthood; (3) Functional glutamatergic innervation of CRH neurons in the hypothalamus is markedly reduced, and (4) this experience-induced reduction of excitatory innervation resolves by adulthood indicating that the diminished excitatory input to the CRH neuron is not required for the maintenance of the ‘re-programmed’ expression levels of this gene; (5) Augmented early-life experience induces a marked increase in levels of NRSF, a transcription factor negatively regulating Crh gene expression, and this repressor binds directly to the regulatory region of the gene; (6) Increased NRSF levels persist to adulthood. Together, these results support the idea that augmented early-life experience reduces excitatory input onto CRH-expressing neurons. This reduced excitation is associated with--and perhaps initiates--a cascade of molecular events, including upregulation of NRSF, which may function to repress CRH expression. Whereas excitation to the CRH neuron normalizes with age, augmented NRSF levels persist, potentially contributing to maintenance of the enduring repression of CRH expression.
Our data indicate that the input organization of neurons in the developing hypothalamus is not hard-wired but, rather, may be influenced by early-life experience. Combining molecular, morphological, ultrastructural and functional approaches, we found that the number of excitatory, vGlut2-immunoreactive boutons contacting CRH neurons was diminished in ‘experience-augmented’ rats. This was associated with reduced vGlut2 levels in the PVN and reduced frequency of mEPSCs, and all these parameters remained unchanged in inhibitory synapses. The decreased frequency of mEPSCs could be interpreted either as reduced presynaptic excitatory elements, or reduced release probability (Regehr et al., 2001); however the reduction of asymmetric synapses onto CRH neurons, observed using electron microscopy (EM), supports the former possibility (the minor but significant changes in the amplitude of the mPSCs suggest postsynaptic changes that will need further investigation). Similarly, reduced total vGlut2 levels might be a result of reduced synapse size rather than synapse number, but the confocal- and EM studies clarified that the number of excitatory synapses onto identified CRH neurons was attenuated after augmented early-life experience. Together, these data provide evidence for significant reduction of excitatory input onto CRH-expressing neurons of experience-augmented rats, i.e., a ‘re-wiring’ of these neurons.
CRH neurons in the PVN integrate excitatory and inhibitory drives from a number of sources. Both stimulatory and inhibitory afferents from limbic and cortical areas are known to coordinate CRH expression and release (Sawchenko and Swanson, 1983), yet there is little direct innervation of CRH neurons from these regions (Herman et al., 2004). Instead, axons originating in hippocampus, septum, amygdala and prefrontal cortex contact local (peri-PVN) or remote (BnST, preoptic area) relay GABAergic or glutamatergic neurons (Tribollet and Dreifuss, 1981; Herman et al., 2004). Monosynaptic GABAergic inputs onto CRH neurons originate in regions surrounding the PVN and the BnST, and maintain a strong inhibitory tone over CRH neurons (Roland and Sawchenko, 1993; Cullinan et al., 1996). Suppression of this tonic GABAergic inhibition enables secretion of CRH in response to glutamatergic activation (Cole and Sawchenko, 2002; Bartanusz et al., 2004), and functional changes in GABAergic synapses abutting CRH neurons has been described in response to physiological challenges (Miklos and Kovacs, 2002; Verkuyl et al., 2004; Ziegler et al., 2005). Glutamatergic inputs onto CRH cells in PVN originate from other hypothalamic nuclei and from BnST and amygdala (Aubry et al., 1996; Kiss et al., 1996; Di et al., 2003; Herman et al., 2004; Wittmann et al., 2005). Indeed, our previous work suggests that augmented early-life experience may reduce such excitatory input by activating a powerful inhibitory signal to amygdala or BnST from the thalamic paraventricular nucleus (Bhatnagar and Dallman, 1998; Fenoglio et al., 2006). The ability of glutamate and GABA to trigger direct synaptic actions in presumed CRH-neurons (Boudaba et al., 1996; Boudaba et al., 1997), and the fact that excitation of these neurons provokes rapid transcription of the Crh gene (Cole and Sawchenko, 2002; Foradori et al., 2007) whereas elevating brain GABA levels represses CRH expression (Tran et al., 1999; Cole and Sawchenko, 2002) support the idea that the reduced balance of excitation/ inhibition found here after early-life experience leads to suppression of CRH expression.
The consequences of early-life experience, i.e., a phenotype of reduced stress-responsiveness (Plotsky and Meaney, 1993; Avishai-Eliner et al., 2001a) and improved learning and memory (Liu et al., 2000; Fenoglio et al., 2005) is associated with life-long increased expression of GR in hippocampus and reduced expression of CRH in PVN neurons (Plotsky and Meaney, 1993; Liu et al., 1997; Fenoglio et al., 2005). We have previously found that suppression of CRH mRNA commenced already on postnatal day 9, and that reduction of CRH-CRH receptor signalling sufficed to endow immature rats with this phenotype (Fenoglio et al., 2005). Therefore, we focused here on the mechanisms by which augmented early-life experience regulated CRH expression, ‘re-programming’ this expression at lower levels. As mentioned above, we found that early-life experience reduced excitation to CRH-expressing neurons and increased expression of the transcriptional repressor NRSF. However, while the experience-induced repression of CRH expression persisted, the innervation of the CRH neurons normalized by adulthood. These findings suggest that reduced excitation might contribute to the initiation of the ‘re-programming’ of CRH expression levels, but was not required for its maintenance. Rather, the cellular programs repressing expression of the Crh gene have become autonomous of the reduced excitatory input.
What might these cellular mechanisms be? The levels of the transcriptional repressor NRSF were dramatically (5-12 fold) elevated in PVN of experience-augmented rats, and this elevation persisted in parallel to repressed CRH expression. NRSF, originally thought to silence neuronal genes in non-neural cells, is now known to be involved in neuronal plasticity (Palm et al., 1998). NRSF binds to a 21-base pair sequence (NRSE), and this sequence is found within the regulatory region (intron) of the Crh gene (Seth and Majzoub, 2001). Our data are the first to show that NRSF binds specifically to this NRSE in vivo, in the hypothalamus of the developing rat. This binding should promote recruitment of co-factors contributing to epigenetic chromatin modification (Zheng et al., 2009), and, if increased after augmented maternal care, should result in repression of transcription of the Crh gene. The involvement of epigenetic mechanisms in the programming of the hypothalamic pituitary adrenal axis by early-life maternal care has been demonstrated at the level of the promoter region of GR (Weaver et al., 2005; 2006; 2007). The enduring enhancement of hypothalamic NRSF levels, combined with the specific binding of this repressor to the Crh gene are consistent with effects of this repressor on transcription of the Crh gene throughout life.
The relationship of reduced excitation and NRSF expression is intriguing. It has been shown that neuronal activity can influence NRSF expression; for example, abnormally increased synchronized excitation (seizures) augments NRSF expression in hippocampus (Palm et al., 1998; McClelland et al., Society for Neuroscience Abstracts, 2008). In addition, and germane to the current study, the levels of NRSF diminish rapidly during the developmental period discussed here (Palm et al., 1998). Whereas the regulation of this developmental decline of NRSF is not fully understood, it is tempting to speculate that age-dependent increase of synaptic input might be responsible for this reduction. If so, than the decreased excitatory synaptic input onto the hypothalamic CRH cell, provoked by augmented maternal care, should result in increased NRSF levels, as found here.
In summary, early-life experience leads to transient re-wiring of hypothalamic neurons and re-programming of Crh gene expression at suppressed levels. These result in a life-long phenotype of reduced stress-responsiveness as well as improved cognitive function. Therefore, understanding the basis of this experience-related plasticity is profoundly important to human health and disease, and should provide the foundation of future therapeutic interventions.
This work was supported by NIH NS28912, MH73136, DK-060711, DK-080000 and DK-070723. The authors thank Bart Pollux, PhD, for statistical advice and Jessica Cope, BSc, for assistance with the in situ hybridization.